parasitic nematodes manipulate plant development to

Parasitic nematodes manipulate plant development toestablish feeding sitesShahid Siddique and Florian MW Grundler

Available online at www.sciencedirect.com

ScienceDirect

Cyst and root-knot nematodes, the two economically most

important groups of plant parasitic nematodes, induce

neoplastic feeding sites in the roots of their host plants. The

formation of feeding sites is accompanied by large-scale

transcriptomic, metabolomic, and structural changes in host

plants. However, the mechanisms that lead to such remarkable

changes have remained poorly understood until recently. Now,

genomic and genetic analyses have greatly enhanced our

understanding of all aspects of plant–nematode interaction.

Here, we review some of the recent advances in understanding

cyst and root-knot nematode parasitism. In particular, we

highlight new findings on the role of plant hormones and small

RNAs in nematode feeding site formation and function. Finally,

we touch on our emerging understanding of the function of

nematode-associated secretions.

Address

Molecular Phytomedicine, INRES, University of Bonn, Karlrobert-

Kreiten-Straße 13, D-53115 Bonn, Germany

Corresponding author: Grundler, Florian MW ([email protected])

Current Opinion in Microbiology 2018, 46:102–108

This review comes from a themed issue on Host microbe interac-

tions: parasitology

Edited by Pascal Maser

For a complete overview see the Issue and the Editorial

Available online 13th October 2018

https://doi.org/10.1016/j.mib.2018.09.004

1369-5274/ã 2018 Elsevier Ltd. All rights reserved.

IntroductionPlant-parasitic nematodes (PPNs) affect almost all major

crops. The presently more than 4100 described PPN

species are estimated to cause over 80 billion USD in

agricultural loss per year [1]. The full extent of worldwide

nematode damage is likely underestimated, particularly

in developing countries, since growers are often unaware

of the presence of these small, soil-borne pathogens.

Additionally, the symptoms caused by PPNs are often

non-specific, making it difficult to attribute crop losses to

nematode damage. The small size, biotrophic life style,

non-synchronized infection, and lack of a reliable trans-

formation method make PPNs difficult experimental

organisms. Studies on the molecular aspects of plant–


nematode interactions have therefore lagged behind

those in other pathosystems.

PPNs use a hollow protrusible stylet to break into the

plant cells, withdraw nutrients, and release both pro-

teinaceous (effectors) and non-proteinaceous mole-

cules. The hollow stylet is connected to three enlarged,

specialized esophageal gland cells, which produce the

effector molecules that are secreted into the host tis-

sues to facilitate parasitism. Each of the three esoph-

ageal glands consists of a single cell that contains an

unusually long cytoplasmic extension ending in an

ampulla. The effector proteins are synthesized in the

gland cell and transported to the ampulla in membrane-

bound granules. The ampulla in turn is connected to

the lumen of the oesophagus by a valve. Some of the

genes encoding oesophageal secretions are likely to

have been acquired from prokaryotic microbes via

horizontal gene transfer [2]. The development of stylet

and esophageal gland cells producing effector mole-

cules are among the most striking adaptations that

enable PPNs to maintain a unique long-term parasitic

relationship with their hosts.

Different species of PPNs feed on a range of plant tissues,

including flowers, stems, leaves, and roots; however, most

species feed on roots. Based on their feeding habits, PPNs

can be broadly categorized as either ectoparasitic or

endoparasitic (Figure 1). In this review, we focus on a

complex and economically devastating group of sedentary

endoparasitic PPNs including cyst nematodes (CNs; Het-erodera spp. and Globodera spp.) and root-knot nematodes

(RKNs; Meloidogyne spp.).

Infective-stage CN and RKN juveniles (J2) invade the

plant root near the tip and move through different

tissue layers to reach the vascular cylinder, where CNs

induce the formation of a syncytium (a multinucleate

fusion of cells resulting from partial cell wall dissolu-

tion) and RKNs induce the formation of 5–7 giant cells

(Figure 2). In the case of RKN, proliferation of the

tissue surrounding the nematode and the giant cells

leads to the formation of a typical gall, which is

observed as a primary symptom of infection. The

establishment of feeding sites (syncytia and giant

cells) enables CNs and RKNs for taking large amounts

of nutrients from the plant, facilitates nematode

growth, and induces a pathologically disturbed alloca-

tion of photosynthetic products that reduces plant

growth and yield.

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Parasitic nematodes establish feeding sites Siddique and Grundler 103

Figure 1

Root-knot nematode

EpidermisEndodermis

Cortex

Pericycle

Cyst nematode

Migratory endoparasitic

Migratory ectoparasitic

Migratory ectoparasitic

Current Opinion in Microbiology

Overview of feeding habits of plant-parasitic nematodes. Plant-parasitic nematodes display a variety of feeding habits and can be broadly

categorized as either ectoparasites or endoparasites. Migratory ectoparasitic nematodes stay vermiform throughout their life cycle and all stages

are capable of feeding on roots of a broad range of host plants. Examples of migratory ectoparasitic nematodes include awl nematodes

(Dolichodorus spp.), sting nematodes (Belonaloaimus spp.), needle nematodes (Longidorus spp.), and dagger nematodes (Xiphinema spp.).

Members of the latter two genera extend periods of feeding at their feeding sites and are able to induce the formation of nurse cell in the root tip.

They also act as vectors of specific plant viruses. Migratory endoparasitic nematodes can cause high yield losses in a variety of field crops. In

addition to the direct damage they inflict on the host, these nematodes promote secondary bacterial and fungal infections. Examples of migratory

endoparasitic nematodes include lesion nematodes (Pratylenchus spp.), burrowing nematodes (Radopholus spp.), and rice root nematodes

(Hirschmanniella spp.). Sedentary endoparasitic plant-parasitic nematodes include cyst nematodes (Heterodera spp. and Globodera spp.) and

root-knot nematodes (Meloidogyne spp.). Both cyst and root-knot nematodes induce hypermetabolic feeding sites in roots, which are the only

source of nutrients for nematodes throughout their life cycle. While the host range of most Meloidogyne species tends to be broad, it remains

rather narrow for most cyst nematode species.

As obligate biotrophs, CN and RKN are entirely-depen-

dent on plant-derived nutrients and solutes to fulfil their

energy requirements throughout their weeks-long life

cycles. Thus, both the syncytium and giant cells have

evolved into a sink tissue that caters to the needs of the

rapidly developing nematode. The cytoplasm of these

feeding sites is dense and contains numerous organelles,

including mitochondria, plastids, ribosomes, the Golgi

apparatus, and the smooth endoplasmic reticulum. Fur-

thermore, the central vacuole in these cells is replaced by

several small vacuoles, and numerous ingrowths are

formed at the cell wall interface with xylem cells, which

are thought to increase the surface area for translocation of

nutrients.

A series of transcriptomic, metabolomic, and proteomic

analyses performed over the last decade showed that the

genes and pathways involved in primary metabolism are

specifically upregulated in both syncytia and giant cells

[3–5]. As previous excellent reviews describe the metab-

olism and functioning of feeding sites [6–8] and discuss


the modulation of plant immunity in response to nema-

tode infection [9,10], we will focus on recent progress in

understanding the formation and functioning of both

types of feeding sites. The first section of our review

explores homeostasis of two crucial plant hormones (cyto-

kinin and gibberellin) that facilitate the formation and

functioning of nematode feeding sites. The second sec-

tion reviews current progress in understanding the role of

small RNAs in syncytium and giant cell formation. The

last section highlights our nascent understanding of nem-

atode-associated secretions that are released into hosts to

facilitate various aspects of parasitism.

Changes in hormone homeostasis during theformation of feeding sitesThe involvement of various plant hormones in plant–

nematode interaction is well-documented, and we refer

readers to previous excellent reviews on the roles of

auxin, salicylic acid, jasmonic acid, and ethylene [6,11].

Research during the last few years has established cyto-

kinins and gibberellins as important players in nematode


104 Host microbe interactions: parasitology

Figure 2

Male

Male

Female

SyncytiumInfective juvenile

Eggs hatchCyst

Gall

Giant cells

Female

Eggmass

Eggs hatchInfective juvenile

Current Opinion in Microbiology

Life cycle of cyst and root-knot nematodes.

(a and b) Second stage infective juveniles of cyst nematodes hatch from the eggs and enter host roots near the tip. They pass the different

tissue layers and invade the vascular cylinder, where they select a single root cell to induce the formation of a syncytium. The juveniles commance

feeding, lose their ability to move and increase in size. Within about two weeks they undergo three moults to reach adulthood. Females become

lemon-shaped and continue feeding for about another two weeks. Female associated syncytia are large and include several hundred root cells.

Male juveniles stop feeding after the third stage; the syncytia remain small. During their consecutive moults they regain a vermiform body shape

and mobility. The adult males hatch from the juvenile cuticle and mate with females. After mating, females produce several hundred eggs which

mostly remain within in the female body. The females die and their body wall turns into a robust cyst harbouring several hundred eggs that may

stay viable in the soil for more than a decade.

(c and d) Second stage infective juveniles of root-knot nematodes hatch and penetrate roots behind the tip. They migrate towards the root tip

meristem where they turn round and enter the vascular cylinder. They cease to migrate in the differentiation zone and select five to seven cells to

induce the formation of giant cells. As soon as they commance feeding, they start to develop, lose their ability of locomotion and increase in size.

Plant tissue surrounding the growing juvenile proliferates and forms a gall – the root-knot. The most important root-knot nematode species

reproduce via parthenogenesis. After moulting three times, the adult female starts to lay eggs into a gelatinous matrix that is secreted on the root

surface and embeds and protects the eggs until the juvenile hatch. N = female nematode, X = xylem, asterisks = syncytial feeding cells (2 A) and

giant cells (2B), arrow heads = remnants of dissolved plant cell walls, scale bar = 20 mm.

feeding site formation, and we will therefore review

recent advances in understanding the role of these two

hormones in CN and RKN parasitism.

One of the first events induced by cyst and root-knot

nematodes upon feeding site development is activation of

the host cell cycle [12]. It is generally assumed that

nematodes manipulate the production of the phytohor-

mone cytokinin and modulate downstream signalling

events to activate cell division [13]. Nematodes have

been shown to produce cytokinin in vitro [14]; however,


whether the hormone is secreted into host plants to

facilitate parasitism was unknown. In three recent pub-

lications, it has been shown that both CNs and RKNs

induce cytokinin signalling at their feeding sites and also

in the neighbouring cells destined to be incorporated into

the feeding sites. Moreover, Arabidopsis plants impaired

in cytokinin biosynthesis or cytokinin signalling are sig-

nificantly less susceptible to infection with both CNs and

RKNs. Cytokinin signalling was also shown to be

required for cell cycle activation at the site of infection,

thus playing a key role in the expansion of both syncytia



Table 1

MicroRNAs (miR) involved in cyst nematode and root-knot

nematode feeding site formation

miRNA Target mRNA Host plant Nematode Ref

miR858 MYB83 Arabidopsis H. schachtii [32��]miR827 NLA Arabidopsis H. schachtii [37]

miR396 GRF1/GRF3 Arabidopsis H. schachtii [38]

miR390 ARFs Arabidopsis M. javanica [25]

miR159 MYB33 Arabidopsis M. incognita [31]

miR319 TCP4 Tomato M. incognita [27]

NLA, Nitrogen Limitation Adaptation; GRF, Growth-Regulating Fac-

tor; ARF, Auxin Response Factors; TCP, Teosinte branched Cycloidea

Proliferating cell nuclear antigen factor.

and giant cells [15��,16�,17]. Notably, CNs not only

produce cytokinin but also release it into infected tissues

to activate the host cell cycle at and around the syncy-

tium. These results clearly showed that cytokinins are

central to feeding site formation and expansion. Intrigu-

ingly, however, mutant plant lines with increased sensi-

tivity to cytokinin are also less susceptible to CNs, due to

heightened immune responses in these plants [17]. Based

on these recent studies, we suggest that a tightly con-

trolled activation of cytokinin is required for optimal

development of both CNs and RKNs.

Although cytokinin seems to play a similar role in ontog-

eny of both syncytia and giant cells, differences have been

identified in the regulation of cytokinin biosynthesis,

catabolism, and signalling genes in response to infection

with CNs and RKNs, suggesting that these two types of

nematode manipulate the cytokinin signalling pathway

differently [16�]. For example, a cytokinin-synthesizing

gene has been identified in CNs and characterised for its

role in parasitism [15��], whereas identification of similar

genes from RKNs has remained elusive. Considering that

much higher levels of cytokinins have been previously

detected in RKNs as compared to CNs [14], however, it is

reasonable to assume that cytokinin-synthesizing genes

are also present in RKN.

Gibberellic acids (GA, or gibberellins) are a large family of

tetracyclic diterpenoids that regulate a variety of growth

and developmental processes in plants. Previous studies

indicated the possible involvement of GA in response to

infection with CNs and RKNs [18–20]. In a recent study,

Yimer and colleagues performed an in-depth characteri-

zation of the role of GA in rice upon infection with a RKN.

They found that RKN infection leads to accumulation of

a specific GA, GA12, at the infection site. Moreover, rice

mutants impaired in GA biosynthesis or signalling were

less susceptible to infection with RKN. Notably, their

detailed functional characterization showed that GA med-

iates susceptibility to nematodes by suppressing defences

regulated by jasmonic acid (JA) during RKN infection

[21��]. Interestingly, GA-induced susceptibility to nema-

todes also depends on auxin transport, as treatment of rice

plants with NPA (N-1-naphthylphthalamic acid), a polar

auxin transport inhibitor, reduced rice susceptibility to

RKN. Jasmonic acid also functions in plant defence

against CNs [22], and it is therefore plausible that GA

might similarly suppress JA-related defences during CN

infection.

The role of microRNAs (miRNAs) in plant–nematode interactionsThe formation of nematode feeding sites is accompanied

by massive transcriptomic changes [4,5]. Until recently,

the details of the mechanisms that lead to such a global

transcriptional shift remained mostly unknown, but

emerging evidence points to the importance of


microRNAs (miRNAs). These small non-coding RNAs

regulate gene expression by binding to their target mes-

senger RNA (mRNA), leading to mRNA degradation,

translational repression, or altered transcriptional activity

[23]. A number of studies have documented changes in

miRNA expression in response to infection by CNs and

RKNs in Arabidopsis [24–26], tomato [27], soybean

[28,29], and cotton [30]. Notably, plant mutants impaired

in miRNA processing have reduced susceptibility to both

CNs and RKNs, indicating a role for miRNAs in feeding

site formation [24,31].

Recent studies have focused on the mechanisms by which

individual miRNA families contribute to transcriptome

reprogramming during the formation of syncytia and giant

cells (Table 1). For example, miR858 has been shown to

play a role in syncytium formation by regulating the

expression of its target transcription factor

MYB83. Constitutive overexpression of miR858 leads to

decreased susceptibility to CNs, while reduced abun-

dance of miR858 leads to increased susceptibility. Simi-

larly, overexpression of a modified version of MYB83 that

cannot be cleaved by miR858 leads to increase suscepti-

bility to CNs. Notably, transcriptome analysis of MYB83overexpression lines showed that 16.6% of the syncytial

transcriptome is regulated by MYB83, indicating a role for

a miR858-MYB83 regulatory system in gene expression

modulation during CN parasitism [32��]. Together, these

different studies make it clear that host miRNA pathways

are powerful targets for nematodes to modulate large-

scale changes in gene expression inside their feeding site.

However, the mechanisms by which nematodes are able

to manipulate the host miRNA expression remain unex-

plored. We speculate that nematodes may release effec-

tors that interfere with the host miRNA biogenesis path-

ways, thus regulating the expression of specific classes of

miRNAs.

Plant use a defence mechanism called host-induced gene

silencing (HIGS) in which small RNAs produced within

the host silence the expression of targeted pathogen or

parasite mRNAs in trans. Plant-based HIGS is effective



against fungi, insects, nematodes, and parasitic plants

[33]. These findings suggest that the exchange of small

RNAs between plants and pathogens/parasites might be a

common feature of plant–pathogen/parasite interactions.

Based on these observations, we speculate that miRNAs

from nematodes may similarly act trans-species to regu-

late large-scale changes in host gene expression.

Nematode effectors at the heart of CN andRKN parasitismThe formation of syncytia and giant cells is facilitated by

the release of a cocktail of proteinaceous (effectors) and

non-proteinaceous secretions inside the host cell. CN and

RKN effectors can be separated into two classes based on

their functions: firstly, suppression of host immune

responses, secondly, formation and functioning of feeding

site. An increasing number of effectors belonging to

either of these two classes have been characterized over

the past few years [34,35]. However, the mechanistic

details by which effectors manipulate host genes and

pathways have been addressed only in a few cases. For

example, a number of nematode effectors have been

shown to suppress host immune responses, but the exact

mechanism by which these effectors achieve such sup-

pression remains unknown. We have summarized the CN

and RKN effectors that have been described during the

last three years in Table 2.

Efforts to predict effectors (proteinaceous secretions)

generally begin with in silico analysis for firstly, the

presence of an N-terminal signal peptide that directs

the protein into the secretory pathway, secondly, the

absence of a transmembrane domain, and thirdly, simi-

larity to available data from other nematode species.

Further characterization of a putative effector often

includes transcript localization by in situ hybridization

and expression profiling during nematode development.

Although these steps may help unravel the putative

Table 2

Cyst nematode and root-knot nematode effectors involved in facilitat

Effector Nematodes Putative function

MjTTL5 M. javanica Immune suppres

MeTCTP M. enterolobii Immune suppres

MgGPP M. graminicola Immune suppres

MiMsp40 M. incognita Immune suppres

HgGLAND18 H. glycines Immune suppres

HsTyr H. schachtii Feeding site form

10A07 H. schachtii Feeding site form

GS G. pallida Feeding site form

MiSGCR1 M. incognita Facilitates early s

MiPFN3 M. incognita Feeding site form

HsPDI H. schachtii Feeding site form

CLE41/44 H. schachtii Feeding site form

Hs30D08 H. schachtii Feeding site form

HsGLAND4 H. schachtii Feeding site form


function of the effector, the actual localization of effectors

inside the host tissues is rarely confirmed (e.g. through

immunolabelling), thus making it difficult to exclude the

possibility that these effectors are never secreted into host

tissues.

Considering that effectors constitute only a minority of

nematode-secreted proteins, additional tools are needed

to establish that a secretory protein is a bona fide effector.

Excitingly, a recent study identified a 6-bp (ATGCCA)

dorsal gland promoter element (DOG box) that is highly-

enriched in the promoter regions of several established

dorsal gland effectors in cyst nematodes [36��]. Further-

more, genes with more DOG boxes in their promoter

regions were more likely to encode a signal peptide for

secretion. The discovery of the DOG box has opened the

door to the development of tools to distinguish effectors

from other secreted proteins. In the future, more such

tools are needed to identify the effectors with a role in

parasitism.

Future perspectivesThe recent progress in understanding plant–nematode

interactions underscores the necessity to elucidate the

integrated molecular framework that explains how nema-

todes are able to form and maintain their unique feeding

sites inside the plants. The research during the last

several years has identified nematode secretions as key

to feeding site formation and maintenance. It is becoming

increasingly clear that nematodes release not only pro-

teinaceous but also non-proteinaceous molecules to

manipulate the host cell machinery. A challenge for

the future will be to establish assays and tools that can

better identify these secretions. It will also be crucial to

develop transformation protocols for PPNs, so that we can

specifically interfere with various aspects of parasitism

and study the consequence of such manipulations on the

infection process.

ing parasitism

Ref.

sion via activation of ROS scavenging [39]

sion via unknown mechanism [40]




ation via unknown mechanism [44]

ation by manipulating post-translational machinery [45]

ation via modulation of redox homeostasis [46]

tages of infection via unknown mechanism [47]

ation via manipulation of actin filaments [48]

ation via modulation of redox homeostasis [49]

ation via vascular stem cell pathway manipulation [50]

ation via interaction with host SMU2 protein [51]

ation via binding to promoter of LTP genes [52�]



Conflict of interest statementNothing declared.

AcknowledgementsWe apologize to all authors whose work could not be cited due to spacelimitations. We gratefully acknowledge Miroslaw Sobczak for providingmicroscopic pictures. Shahid Siddique was supported by grants fromGerman Research Foundation (SI 1739/3-1 and SI 1739/5-1).

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28. Tian B, Wang SC, Todd TC, Johnson CD, Tang GL, Trick HN:Genome-wide identification of soybean microRNA responsiveto soybean cyst nematodes infection by deep sequencing.BMC Genomics 2017, 18:572.


http://refhub.elsevier.com/S1369-5274(18)30036-5/sbref0005

















































































































29. Li XY, Wang X, Zhang SP, Liu DW, Duan YX, Dong W:Identification of soybean microRNAs involved in soybean cystnematode infection by deep sequencing. PLoS One 2012, 7:e39650.

30. Pan X, Nicholas RL, Li C, Zhang B: MicroRNA-target generesponses to root-knot nematode (Meloidogyne incognita)infection in cotton. Genomics 2018 http://dx.doi.org/10.1016/j.ygeno.2018.02.013.

31. Medina C, da Rocha M, Magliano M, Ratpopoulo A, Revel B,Marteu N, Magnone V, Lebrigand K, Cabrera J, Barcala M et al.:Characterization of microRNAs from Arabidopsis gallshighlights a role for miR159 in the plant response to the root-knot nematode Meloidogyne incognita. New Phytol 2017,216:882-896.

32.��

Piya S, Kihm C, Rice JH, Baum TJ, Hewezi T: Cooperativeregulatory functions of miR858 and MYB83 during cystnematode parasitism. Plant Physiol 2017, 174:1897-1912.

This study demonstrated that ArabidopsismiRNA858 (miR858) and itstarget transcription factor MYB83 are involved in large-scale transcrip-tome programming of syncytium induced by cyst nematodes.

33. Weiberg A, Jin HL: Small RNAs - the secret agents in the plant-pathogen interactions. Curr Opin Plant Biol 2015, 26:87-94.

34. Ali MA, Azeem F, Li HJ, Bohlmann H: Smart parasitic nematodesuse multifaceted strategies to parasitize plants. Front Plant Sci2017, 8:1699.

35. Juvale PS, Baum TJ: "Cyst-ained" research into Heteroderaparasitism. PLoS Pathog 2018, 14:e1006791.

36.��

Eves-van den Akker S, Laetsch DR, Thorpe P, Lilley CJ,Danchin EGJ, Da Rocha M, Rancurel C, Holroyd NE, Cotton JA,Szitenberg A et al.: The genome of the yellow potato cystnematode, Globodera rostochiensis, reveals insights into thebasis of parasitism and virulence. Genome Biol 2016, 17:124.

This study identified a 6-bp (ATGCCA) dorsal gland promoter element thatis enriched in the promoter regions of several well-established dorsalgland effectors in cyst nematodes. Furthermore, genes with more DOGboxes in their promoter regions were more likely to encode a signalpeptide for secretion.

37. Hewezi T, Piya S, Qi MS, Balasubramaniam M, Rice JH, Baum TJ:Arabidopsis miR827 mediates post-transcriptional genesilencing of its ubiquitin E3 ligase target gene in the syncytiumof the cyst nematode Heterodera schachtii to enhancesusceptibility. Plant J 2016, 88:179-192.

38. Hewezi T, Maier TR, Nettleton D, Baum TJ: The ArabidopsismicroRNA396-GRF1/GRF3 regulatory module acts as adevelopmental regulator in the reprogramming of root cellsduring cyst nematode infection. Plant Physiol 2012, 159:321-335.

39. Lin BR, Zhuo K, Chen SY, Hu LL, Sun LH, Wang XH, Zhang LH,Liao JL: A novel nematode effector suppresses plant immunityby activating host reactive oxygen species-scavengingsystem. New Phytol 2016, 209:1159-1173.

40. Zhuo K, Chen JS, Lin BR, Wang J, Sun FX, Hu LL, Liao JL: A novelMeloidogyne enterolobii effector MeTCTP promotesparasitism by suppressing programmed cell death in hostplants. Mol Plant Pathol 2017, 18:45-54.


41. Chen JS, Lin BR, Huang QL, Hu LL, Zhuo K, Liao JL: A novelMeloidogyne graminicola effector, MgGPP, is secreted intohost cells and undergoes glycosylation in concert withproteolysis to suppress plant defenses and promoteparasitism. PLoS Pathog 2017, 13:e1006301.

42. Niu JH, Liu P, Liu Q, Chen CL, Guo QX, JunmeiYin, GuangsuiYang,Jian H: Msp40 effector of root-knot nematode manipulatesplant immunity to facilitate parasitism. Sci Rep 2016, 6:19443.

43. Noon JB, Qi MS, Sill DN, Muppirala U, Eves-van den Akker S,Maier TR, Dobbs D, Mitchum MG, Hewezi T, Baum TJ: APlasmodium-like virulence effector of the soybean cystnematode suppresses plant innate immunity. New Phytol 2016,212:444-460.

44. Habash SS, Radakovic ZS, Vankova R, Siddique S, Dobrev P,Gleason C, Grundler FMW, Elashry A: Heterodera schachtiiTyrosinase-like protein - a novel nematode effectormodulating plant hormone homeostasis. Sci Rep 2017, 7:6874.

45. Hewezi T, Juvale PS, Piya S, Maier TR, Rambani A, Rice JH,Mitchum MG, Davis EL, Hussey RS, Baum TJ: The CystNematode effector protein 10A targets and recruits hostposttranslational machinery to mediate its nuclear traffickingand to promote parasitism in Arabidopsis. Plant Cell 2015,27:891-907.

46. Lilley CJ, Maqbool A, Wu D, Yusup HB, Jones LM, Birch PRJ,Banfield MJ, Urwin PE, Akker SE-vd: Effector gene birth in plantparasitic nematodes: neofunctionalization of a housekeepingglutathione synthetase gene. PLoS Genet 2018, 14:e1007310.

47. Nguyen CN, Perfus-Barbeoch L, Quentin M, Zhao JL, Magliano M,Marteu N, Da Rocha M, Nottet N, Abad P, Favery B: A root-knotnematode small glycine and cysteine-rich secreted effector,MiSGCR1, is involved in plant parasitism. New Phytol 2018,217:687-699.

48. Leelarasamee N, Zhang L, Gleason C: The root-knot nematodeeffector MiPFN3 disrupts plant actin filaments and promotesparasitism. PLOS Pathog 2018, 14:e1006947.

49. Habash SS, Sobczak M, Siddique S, Voigt B, Elashry A,Grundler FMW: Identification and characterization of a putativeprotein disulfide isomerase (HsPDI) as an alleged effector ofHeterodera schachtii. Sci Rep 2017, 7:13536.

50. Guo X, Wang J, Gardner M, Fukuda H, Kondo Y, Etchells JP et al.:Identification of cyst nematode B-type CLE peptides andmodulation of the vascular stem cell pathway for feeding cellformation. PLoS Pathog 2017, 13:e1006142.

51. Verma A, Lee C, Morriss S, Odu F, Kenning C, Rizzo N,Spollen WG, Lin M, McRae AG, Givan SA et al.: The novel cystnematode effector protein 30D targets host nuclear functionsto alter gene expression in feeding sites. New Phytol 2018http://dx.doi.org/10.1111/nph.15179.

52.�

Barnes SN, Wram CL, Mitchum MG, Baum TJ: The plant-parasitic cyst nematode effector GLAND4 is a DNA-bindingprotein. Mol Plant Pathol 2018 http://dx.doi.org/10.1111/mpp.12697.

This study identified cyst nematodes GLAND4 as the first DNA-bindingplant-parasitic nematode effector.






http://dx.doi.org/10.1016/j.ygeno.2018.02.013

http://dx.doi.org/10.1016/j.ygeno.2018.02.013



















































































http://dx.doi.org/10.1111/nph.15179

http://dx.doi.org/10.1111/mpp.12697

http://dx.doi.org/10.1111/mpp.12697

parasitic nematodes manipulate plant development to

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